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Pumpless microflow cytometry enabled by viscosity modulation and immunobead labeling Byeongyeon Kim, Sein Oh, Suyeon Shin, Sang-Gu Yim, Seung Yun Yang, Young Ki Hahn, and Sungyoung Choi Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.8b01804 • Publication Date (Web): 06 Jun 2018 Downloaded from http://pubs.acs.org on June 6, 2018

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Analytical Chemistry

Pumpless microflow cytometry enabled by viscosity modulation and immunobead labeling Byeongyeon Kim,† Sein Oh,† Suyeon Shin,† Sang-Gu Yim,‡ Seung Yun Yang,‡, Young Ki Hahn,*,ǀǀ and Sungyoung Choi*,† †

Department of Biomedical Engineering, Kyung Hee University, Yongin-si, Gyeonggi-do 17104, Republic of Korea

‡

Department of Biomaterials Science, Life and Industry Convergence Institute, Pusan National University, 1268-50 Samrangjin-ro, Miryang 50463, Republic of Korea ǀǀ

Department of New Biology, Daegu Gyeongbuk Institute of Science & Technology (DGIST), Daegu 42988, Republic of Korea

ABSTRACT: Major challenges of miniaturizing flow cytometry include obviating the need for bulky, expensive, and complex pump-based fluidic and laser-based optical systems, while retaining the ability to detect target cells based on their unique surface receptors. We addressed these critical challenges by (i) using a viscous liquid additive to control flow rate passively, without external pumping equipment, and (ii) adopting an immunobead assay that can be quantified with a portable fluorescence cell counter based on a blue light-emitting diode. Such novel features enable pumpless microflow cytometry (pFC) analysis by simply dropping a sample solution onto the inlet reservoir of a disposable cell-counting chamber. With our pFC platform, we achieved reliable cell counting over a dynamic range of 9 to 298 cells/µL. We demonstrated the practical utility of the platform by identifying a type of cancer cell based on CD326, the epithelial cell adhesion molecule. This portable microflow cytometry platform can be applied generally to a range of cell types using immunobeads labeled with specific antibodies, thus making it valuable for cell-based and point-of-care diagnostics.

Counting cells and identifying their phenotypic differences are of critical importance to a range of biomedical applications from in vitro diagnostics to examination of abnormal cells1-3 to cellular engineering to study of cell proliferation and differentiation4,5. For instance, white blood cell count is an important indicator for diagnosing various inflammatory and infectious diseases1. Detection methods based on cell surface receptors are an indispensable tool for identifying target cells from heterogeneous cell populations5. As a fundamental tool for such applications, fluorescence microscopy can provide highresolution cell analysis6,7, but suffers from limited throughput. Although flow cytometry enables highthroughput analysis by detecting individual cells flowing in a single-file stream8,9, the requirement for expensive and sophisticated equipment makes the technology unsuitable in resource-limited settings. Alternatively, imaging flow cytometry allows both qualitative and quantitative analyses by combining the features of fluorescence microscopy and flow cytometry10,11 but partly shares their disadvantages with regard to system size and cost. Over the last decade, a major technical advance in the field of flow cytometry was the incorporation of microfluidics into flow cytometry, which provided unique functionalities such as three-dimensional cell focusing12-15 and on-chip cell sorting16-18. These features can simplify complex fluidic structures used in conventional flow cytome-

try, and allow for high-purity and high-viability cell sorting, while maintaining the throughput performance of conventional flow cytometry. However, most microfluidic counterparts still use the conventional design of flow cytometry, consisting of external pumps and control systems to achieve high flow stability and precision. Thus, their routine use in practical applications can be limited, considering that the additional fluidic accessories significantly increase operational complexity and space. Another advancement was the development of simple detection schemes, which allow for easy-to-use and inexpensive flow cytometric analysis19-23. A smartphone, in conjunction with an add-on microfluidic device that is working as an optofluidic waveguide19, provides the promise of rapid diagnostic testing and automated data analysis. Lens-free imaging platforms enable cost-effective cell counting in resource-limited settings by obviating the need for bulky optics and instead incorporating an algorithm for holographic image reconstruction20,21. Electrical detection methods have been advanced for label-free cell counting by capturing target cells in an immuno-affinity chamber and counting the remaining cells based on the Coulter principle22,23. However, these approaches still rely on bulky and expensive pumping equipment for flow control19-23. In addition, some of them lack the ability to detect target cells based on their specific surface receptors, an essential function of flow cytometry19,20.

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Figure 1. Pumpless microflow cytometry (pFC). (a) Schematic for pumpless cell counting and analysis. Cell samples are delivered into a disposable cell counting chamber by pipetting. The fluorescence characteristics of flowing cells can be accurately measured without image distortion by passive flow-rate control using a viscous liquid additive. Target cells labeled with immunobeads are then analyzed based on their fluorescence characteristics. The insets show the fluorescence images of breast cancer cells labeled with submicron fluorescent beads taken with the pFC. Scale bar, 50 μm. (b) The compact cell counter consists of a blue light-emitting diode, a ball lens, optical filters, an objective lens, and a 3D-printed sample mount. The cell counter is used to record fluorescence movies during capillary sample loading. Scale bar, 2 cm. (c) The recorded movies were analyzed using a custom counting algorithm.

Here, we present a comprehensive strategy for pumpless microflow cytometry (pFC) analysis that combines passive flow-rate control by a viscous liquid additive and simple immunophenotyping by immunobead labeling (Figure 1). The viscous liquid additive passively controls the velocity of cells within a detectable range during capillary sample loading, thereby eliminating the need for bulky and expensive pumping equipment. Submicron fluorescent beads bound to target cells generate intense fluorescence signals distinguishable from the weaker fluorescence signal of cytoplasmic staining and can be quantified with a portable fluorescence cell counter based on a blue light-emitting diode (LED). These unique features enable microflow cytometric analysis using a disposable cell-counting chamber and the LED-based cell counter, thereby easily converting the counting chamber into a pFC. We demonstrate the capability of the pFC for reliable cell counting by measuring cell counts as low as 8.9 cells/µL. We then use the pFC to detect the immunophenotypic differences between two different types of breast cancer cells for CD326, the epithelial cell adhesion molecule. With these results, we demonstrate that the integrated approach offers a new possibility of truly portable and disposable flow cytometric analysis.

EXPERIMENTAL SECTION pFC design and fabrication. The pFC is made up of two main systems: fluidics and optics. The fluidic system is a commercial cell-counting chamber (100 µm in height) purchased from Logos Biosystems, Inc. (Korea) and is used to deliver cells to an interrogation area for fluorescence detection. Instead of using bulky and expensive pumping equipment as in conventional flow cytometry and its microfluidic counterparts, the pFC regulates flow rate under a threshold value using a viscous liquid additive, polyvinylpyrrolidone (PVP; Sigma Aldrich, USA). The molar mass of the PVP used in this study was 360,000 g/mol. The optical system is composed of a blue LED (457 nm; LED Engin, Inc., USA) to illuminate fluorescent particles and cells, optical filters with 474 nm and 525 nm center wavelength (Semrock, Inc., USA), a 10× objective lens (Nikon Corp., Japan), and a CMOS sensor (FLIR, Inc., Canada). These optical components were compactly assembled in a housing fabricated using a 3D printer (DWS Systems Corp., Italy) as previously described.24 For pFC analysis, cells were immuno-labeled with submicron fluorescent particles and then delivered into the counting chamber by simple pipetting. During capillary sample


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loading, fluorescence movies were recorded using the CMOS sensor connected to a laptop and were analyzed using a custom MATLAB (The MathWorks, Inc., USA) program developed to automatically count cells and quantify their fluorescence characteristics. The low weight (250 g) and small size (257 cm3) of the pFC allow for portable flow cytometric analysis. Sample preparation. Three cell lines (K562, MCF7, and MDA-MB-231) were cultured in Roswell Park Memorial Institute (RPMI) 1640 medium (Welgene, Inc., Korea) supplemented with 10% (v/v) fetal bovine serum (Welgene) and 1% antibiotic solution containing penicillin and streptomycin at 37 °C in a humidified atmosphere containing 5% CO2. Before experiments, these three cell lines were washed with Dulbecco's Phosphate-Buffered Saline (DPBS) solution supplemented with 1% bovine serum albumin. K562 cells were used for cell counting application and were prepared by staining the cells with a nucleic-acid stain, SYTO 13 (Thermo Fisher Scientific, Inc., USA) at a concentration of 5 μM, and then washing them with DPBS. MCF7 and MDA-MB-231 are representative breast cancer cell lines expressing different levels of CD3262,25 and were used for demonstration of the ability of the pFC to identify different cell types based on cell surface-receptor expression. For this application, we performed an immunobead assay. For labeling of fluorescent submicron particles on the cell surface, streptavidincoated fluorescent particles (200 nm in nominal diameter, Spherotech, Inc., USA) were incubated with biotinylated CD326 antibody (R&D Systems, Inc., USA) overnight. After washing with DPBS, 0.2 mg/mL of the antibodyconjugated particles were mixed with cells (1.0 × 106 cells/mL) for 2 h, and then unbound particles were removed by centrifugation. After bead incubation, the cells were stained with carboxyfluorescein diacetate succinimidyl ester, a cell-permeable cytoplasmic staining dye, at a concentration of 1 μM, thereby fluorescently labeling the whole cytoplasmic area of single cells. Due to the high fluorescence intensity of the submicron particles, immuno-bead labeling was clearly distinguished from cytoplasmic staining. Fluorescent polystyrene particles of 6 and 15 μm in diameter were purchased from Polysciences, Inc. (USA) and Invitrogen Corp. (USA), to test the effect of particle size on the counting performance of the pFC. Polystyrene particles of 10 μm in diameter (Polysciences) were used for PVP characterization. Sample analysis. Flow trajectories and fluorescence characteristics of microparticles (15 μm in diameter) were observed using an inverted microscope (Nikon) equipped with a high-speed camera (Vision Research, Inc., USA) or using the pFC and were then analyzed using ImageJ software (National Institutes of Health, USA). The viscosities of the PVP solutions were measured using a 3D-printed capillary circuit as previously reported26. Briefly, the capillary circuit was designed to compare two different liquids, one with known viscosity and the other with unknown viscosity. The unknown viscosity was determined by the volume ratio of two liquids dispensed under the same pressure condition. Cell viability was assessed by trypan

blue and propidium iodide (PI) staining. We note that dead cells were not stained properly in the trypan blue exclusion assay (Figure S1). On the other hand, PI successfully stained dead cells. These results imply that the viabilities of cells suspended in PVP solutions can be overestimated as assessed by trypan blue. In addition, as determined by PI staining, we found that excessive mixing to mix highly viscous solutions supplemented with PVP could generate high shear stresses and affect cell viability (Figure S1). To reduce shear stress and mix the viscous solutions without affecting cell viability, we widened the cross-section of a micropipette tip from ≈0.20 mm2 to ≈1.37 mm2 by cutting its narrow end and confirmed that the viability of the cells was not affected by the high concentration (5%) of PVP for up to 24 h (Figure S2). Flow cytometric analysis on CD326 expression was performed using a BD Accuri C6 flow cytometer (BD Biosciences, USA). For the analysis, the breast cancer cells were stained with anti-CD326 antibodies (BD Biosciences), washed twice, and re-suspended in DPBS. For scanning electron microscope (SEM) imaging, the labeled cells were fixed in 4% paraformaldehyde (Electron Microscopy Sciences, USA), dropped onto a glass slide, and dried at room temperature for 6 h. The morphology of the cells was visualized using a field-emission scanning electron microscope (Hitachi, Japan) at an accelerating voltage of 10 kV after Pt coating with an ion sputter (Hitachi) for 30 s. All the cell counting and analysis experiments were repeated at least three times. Cell counting algorithm. For automatic cell counting, the image frames of a fluorescence movie taken with the pFC were sequentially analyzed with a custom MATALB code. In the counting algorithm, cells were identified and counted from detected contours when both contour size and fluorescence intensity were larger than threshold values. For the immunophenotyping application, the pixel intensity information of each identified cell was stored and used for calculating the mean fluorescence intensity of each cell and the pixel count over a threshold fluorescence value. The code for automatic cell counting is provided in the section of Supplementary Code.

RESULTS AND DISCUSSION Chamber filling dynamics modulated by viscosity. The first requirement to convert a disposable cell-counting chamber into a pFC was to decelerate the filling speed of the chamber with a drop of a sample solution. For 10 µL of a PBS solution dropped onto the inlet reservoir of the counting chamber, the filling speed reached 28.1 mm/s (n = 20 particles), at which speed the suspended particles were too fast to be imaged without an expensive highspeed camera. To address this challenge, we implemented viscosity modulation to decrease the speed of the cells flowing through the chamber and thus accurately detect them using the pFC with a moderate image acquisition rate (100 frames per second). For viscosity modulation, we used a viscoelastic polymer, PVP, which could generate a wide range of viscosity when supplemented in different concentrations (Figure 2a) and which exhibited high bio-



compatibility27. Even at a high PVP concentration (CPVP) of 5 wt%, there was no significant difference between the viabilities of PVP-treated cells and non-treated controls over 24 h (Figure S2). The dynamics of chamber filling is mainly based on capillary pressure (ΔPc) and can be described by rearranging the Hagen-Poiseuille equation.28

∆

,

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an interval of 0.1 s and at ≈0.4 s after sample injection. The micrographs were obtained at 400 fps using a high-speed camera mounted on a microscope. The arrows indicate the start and end positions of the microparticles. Scale bars, 200 µm.

(1)

where u is the velocity of the liquid filling the chamber, h is the chamber height, µ is the Newtonian liquid viscosity, and L is the length of the liquid column filling the chamber.

∆ 2 cos ,

(2)

where σ is the surface tension of the liquid, θ is the contact angle, and w is the chamber width. The viscosity dependence of u enables accurate regulation of particle velocity under a critical velocity of ≈1.5 mm/s at Cpvp = 5 wt% (Figure 2b) such that individual particles can be identified using the pFC without image distortion (Figure 3). These results demonstrated that the liquid additive can act as a fluidic low-pass filter to regulate cell velocity lower than the cutoff velocity, thereby enabling pumpless operation of the pFC.

Figure 3. pFC characterization with 15-μm fluorescent microparticles. (a) Microscopic snapshots from particle-loading movies taken with the pFC. Scale bars, 300 µm. The images were taken right after sample loading. The flow direction is from the left to the right for each image. (b, c) Effect of PVP concentrations on the aspect ratio (b) and peak fluorescence intensity (c) of particle streak-lines (n = 20 particles for each data point). During complete sample loading of ≈60 s, the fluorescence intensity and aspect ratio at CPVP = 5 wt% were maintained without significant variation. The aspect ratio of a particle streak-line is defined as its length-to-width ratio. Figure 2. Viscosity modulation by a viscoelastic polymer, PVP. (a) Viscosities of PVP solutions over a range of concentrations of 0 to 5 wt% (n = 3). (b) Viscosity-dependent and time-varying characteristics of particle velocity at which microparticles (10 µm in diameter) were introduced into the cell counting chamber (n = 8 particles). Each image of the top panels shows the overlay of two micrographs captured with

pFC Characterization. The second requirement to develop the pFC was to build a compact and portable fluorescence cell counter as the optical part of the pFC, making it suitable for the immunobead assay. The field-ofview and corresponding spatial resolution of the imaging cell counter were set to 1.8 mm2 and 1.7 µm, respectively, to identify accurately individual cells while simultaneous-


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ly measuring as many cells as possible. We tested the capability of the cell counter to detect all microscopic particles flowing through its interrogation area without image distortion in combination with the viscosity modulation method. Figure 3 shows how captured images of fluorescent microparticles (15 µm in diameter), serving as a model of cells immuno-labeled with fluorescent submicron particles are affected by CPVP and time after sample loading. As a sample fluid dropped onto the inlet reservoir moves toward the outlet, u rapidly decreases with time (Figure 2b), which can lead to continuous variation in fluorescent and morphological cellular analysis. However, the variation in u significantly decreased at Cpvp = 5 wt% (Figure 2b) due to high viscosity, thereby ensuring robust analysis of the fluorescence intensity and shape of the particles (Figure 3). As CPVP increased to 5 wt%, the aspect ratio of the particle streak-lines remained constant at 1.04 on average (n = 200 particles) during the whole process of sample loading (Figure 3b), which enabled particle image acquisition without motion blur. In addition, the standard deviations of the peak fluorescence intensity (SFL) and aspect ratio (SAR) of the particle streak-lines were improved from SFL = 28.61 and SAR = 0.48 at CPVP = 1% to SFL = 16.94 and SAR = 0.15 at CPVP = 5%. At CPVP = 5%, both the fluorescence intensity and aspect ratio were maintained without significant changes (Figures 3b and 3c), allowing for accurate fluorescent and morphological analyses of individual cells even under unsteady flow conditions. Collectively, the PVP concentration of 5 wt% was found to optimize accurate fluorescent-particle imaging with the pFC. Pumpless cell counting. We applied the optimized pFC protocol to pumpless cell counting. To validate the cell counting ability of the pFC, K562 cells were stained with a nucleic-acid stain, and then mixed with 10 wt% PVP in a 1:1 volume ratio before sample loading into the counting chamber. Titration experiments were prepared by diluting a defined cell suspension (≈3.0 × 105 cells/mL) which was measured with a conventional hemocytometer. For pFC cell counting, 10 µL of sample solution was loaded into the counting chamber by pipetting (Movie S1). During the capillary sample loading, up to 1,254 cells were counted with the dynamic range of cell counting spanning over 2 orders of magnitude. As shown in Figure 4, the cell counts measured by the pFC closely matched the expected cell counts, showing a strong linear relationship with a regression slope of 1.03 and an R2 of 0.9996. Reliable cell counting (CV ≤ 15%) with a conventional hemocytometer required more than 95.0 cells/µL, where CV is the coefficient of variation. Below that concentration, the CV exceeded 16.1%, while the pFC required as low as 8.9 cells/µL with the CV of 9.0%, demonstrating the capability of the pFC for accurate cell counting. This is attributed to the larger volume of sample tested (≈4 µL for pFC and 0.4 µL for hemocytometer) and the larger number of cells counted (≈34.5 cells for pFC and ≈3.5 cells for hemocytometer at the estimated count of 8.9 cells/µL). We note that the maximum instantaneous counting throughput reached up to ≈154 cells/s, while the corresponding aver-

age throughput decreased to ≈2 cells/s due to the significant decrease in u during sample loading at 298 cells/µL. Throughput improvement can be further achieved by incorporating a capillary pump composed of microposts29 at the outlet reservoir of the counting chamber and thus maintaining a constant u during sample loading. Another option to improve the throughput capability of the pFC is to use a camera with a higher frame rate and enable pumpless operation of the pFC at a lower Cpvp and higher u. The detection limit can also be improved by lengthening the chamber and analyzing a larger sample volume. The size of particles and cells is an important parameter for microfluidic particle and cell analysis. Thus, we tested whether particle size affects the pFC performance. At a fixed particle concentration (≈48.2 particles/μL for 6-μm particles and ≈42.7 particles/μL for 15-μm particles), the counting results from the pFC and conventional hemocytometer were closely matched for both particles (Figure S3), demonstrating that cells and particles can be accurately counted by fluorescence using the pFC without any significant effect of the particle size. However, the current form of the pFC cannot be used for particle and cell size analysis, due to the higher channel height than the depth of field of the cell counter. Out-of-focus particles can be estimated to be larger than their actual size. In future work, we aim to develop a specific cell-counting chip that has a lower chamber height and a miniature pressure source, thereby enabling accurate size analysis and generating steady flow conditions.

Figure 4. Pumpless cell counting by the pFC. (a) Microscopic snapshots from cell-loading movies taken using the pFC. Each image was taken ≈3 s after sample loading. The flow direction is from the left to the right of each image. Scale bars, 300 µm. (b) Comparison between the cell counting re-



sults obtained with the pFC and a conventional hemocytometer (n = 4). Even at the low cell concentration (8.9 cells/µL), the pFC provides reliable cell counts (CV = 9.0%), while the hemocytometer results in poor precision (CV = 36.9%). The dotted line represents the unity slope.

Immunofluorescence-based microflow-cytometric analysis. We tested the practical utility of the pFC by performing microflow cytometric analysis of different cell types based on cell surface-receptor expression. Breast cancer cell lines (MCF7 and MDA-MB-231) were used as a model system, which respectively exhibit high and low expression levels for CD326 (Figure S4)2,25. As labeled with the submicron beads conjugated with CD326 antibodies, MCF7 cells were covered with the beads, while MDA-MB231 cells were sparsely coated with the beads, clearly showing the difference in CD326 expression level (Figure 5a). Both cells were also stained with a cytoplasmic staining dye to visualize even non-target cells under fluorescence, and then separately analyzed using the pFC. The submicron beads bound to the target cells (MCF7) generated intense fluorescence signals, which were significantly higher than the cytoplasmic staining of the cells (Figure 5b). Such a phenotypic difference was successfully distinguished with the pFC in the data set (more than 1,041 counts for each cell type), by displaying the measurement parameters for the mean fluorescence intensity of each cell and Cpx that denotes a value by adding 2 to the count of pixels over a threshold fluorescence value (110 a.u.) and ranges from 2 (no bead binding) to 270 (complete bead binding) (Figure 5c and Movie S2). Cpx was mainly used to differentiate the labeled beads and cytoplasmic staining based on fluorescence intensity. Both CD326-positive and -negative cells were visible in fluorescence images obtained using the pFC, but only target cells labeled with immuno-beads were highly visible (Figure 5b). Thus, individual cells labeled with the beads could be clearly identified in the fluorescence-intensity heat map, demonstrating that the pFC in combination with the simple immunobead assay was suitable for detecting target cells based on their specific surface receptor. When gated with Cpx = 2.5, the pFC analyses on the cells matched well with conventional flow cytometry (Figures 5c and S4). There were 2.7 ± 1.7% and 92.2 ± 3.3% CD326-positive cells for MDA-MB-231 and MCF7 measured by the pFC, respectively (n ≥ 4), and there were 4.7% and 98.6% CD326-positive cells for MDA-MB-231 and MCF7 measured by conventional flow cytometry. In-focus imaging is important to obtain robust analysis results in imaging flow cytometry. Although out-of-focus cells can generate weaker fluorescence signals than in-focus cells, the differential cell identification by the pFC could not be affected by a minute difference in focal planes due to the intensive florescence signals generated from the immunobeads bound to the target cells (Figure 5b). Thus, the target cells even in outof-focus planes can be clearly distinguished from the nontarget cells producing weaker fluorescence signals. Although the present pFC provides the capability of analyzing a single phenotypic parameter, multi-parametric analysis can be further achieved using labeling beads in

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various colors and sizes. Since CD326 is a potential marker of cancer stem cells30, the pFC can be directly applicable to rapid analysis of heterogeneous circulating tumor cell populations.31 The present work focuses on demonstrating the pFC platform for cancer cell identification, but our platform is not limited to that specific application. With the availability of a range of antibodies and fluorescent particles, our platform can be easily extended to other applications that require identification of a specific cell population and recognition of phenotypic aberrancies.

Figure 5. Pumpless microflow cytometric analysis of breast cancer cells labeled with immunobeads. (a) SEM images showing the difference in CD326 expression level between two different types of breast cancer cells. Submicron beads (false-colored in green) conjugated with CD326 antibodies are attached to cell surfaces (false-colored in orange). Scale bars, 2 µm. (b) Microscopic snapshots from movies of cellloading, taken with the pFC and their corresponding heat maps. Scale bars, 300 µm. (Enlarged views) The CD326positive cell type (MCF7) exhibits significantly higher fluorescence intensity than the negative cell type (MDA-MB-231).


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The corresponding heat maps also show the difference in fluorescence area over a threshold value (110 a.u.) according to cell type. Scale bars, 20 µm. The flow direction is from the left to the right for each image. (c) Scatter plot of the mean fluorescence intensity and Cpx, a value by adding 2 to the count of pixels over a threshold fluorescence value (110 a.u.) (More than 1,041 data points for each cell type were collected from more than four independent experiments).

CONCLUSIONS In summary, we developed a new type of microflow cytometry that integrates two key technologies toward enabling microflow cytometric analysis in an easy-to-use, simple-to-fabricate, and field-portable manner. The fluidic and optical parts of the pFC were greatly simplified by the passive flow-rate regulation based on the viscous liquid additive and by the immunobead assay that can be tested using the compact and portable cell counter. Thus, dropping a sample solution onto the inlet reservoir of a cell-counting chamber is all the operation required for pFC analysis. With these unique features, we demonstrated the ability of the pFC to detect CD326-positive and negative cells, which opens up a new possibility of simplified microflow cytometric analysis and cell-based diagnostics. Based on the advantages we have demonstrated, our platform can facilitate the widespread use of flow cytometric immunophenopying as an indispensable tool for diagnosing and monitoring clinical conditions.

ASSOCIATED CONTENT Supporting Information The Supporting Information is available free of charge on the ACS Publications website. Supporting figures for comparison of cell viabilities assessed by trypan blue and propidium iodide, the effect of the effect of PVP on cell viability, the effect of particle size on pFC performance, and conventional flow cytometric analysis of breast cancer cells, and supplementary MATLAB script for automatic cell counting (PDF). Supporting video 1 showing the simple sample-dropping process for pumpless flow cytometric analysis (WMV). Supporting video 2 showing the MATLAB-based automatic cell counting program for counting cells and analyzing their fluorescence characteristics. This video was encoded with 16× playback speed (WMV).

AUTHOR INFORMATION Corresponding Author * E-mail: [email protected]. * E-mail: [email protected].

Notes The authors declare no competing financial interest.

ACKNOWLEDGMENT This research was supported by a grant from the Pioneer Research Center Program through the National Research Foundation (NRF) of Korea funded by the Ministry of Sci-

ence, ICT & Future Planning (MSIP) (2013M3C1A3064777), a NRF grant funded by the Korea government (MSIP) (2015R1C1A1A01053990), and a NRF grant funded by the Korea government (MSIP) (2016R1A5A1010148). We thank D. You for assistance in carrying out the experimental work.

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ToC artwork



Figure 1 192x157mm (300 x 300 DPI)


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Figure 4 140x201mm (300 x 300 DPI)



Figure 5 150x277mm (300 x 300 DPI)


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Pumpless microflow cytometry enabled by viscosity ... - ACS Publications

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